2019 - My Year In Spiders

In 2019 I recorded a total of 354 individual spiders representing 103 species, all from VC55 (Leicestershire and Rutland), 29% of the County list. Figure 1 shows where and when I recorded. This year I tended to roam further afield in summer and stuck closer to home in winter:

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Figure 2 scratches beneath the surface to reveal more about these patterns:

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Panels A and C largely represent the expected phenology of spiders, particularly the presence of mature adults which can be identified. It's worth noting that February, although a natural low point in the phenology, is artificially depressed because other commitments meant I was not able to do much field work at this time. October is a little surprising. We have had a warm autumn which means the traditional phenology has been extended, but the heavy rains from mid-September onwards (the wettest ever autumn in the East Midlands) put a big dent in recording effort with roads flooded and some sites too dangerous or difficult to reach. Panels B and D explain much of the pattern seen in panel A. Visits per Month normalizes for recording effort and also reflects the impact of autumn flooding. However, the autumn rising trend of Species per Visit noteworthy - this points to a mild autumn with few frosts.

Target for 2019? I'm thinking at least 500 specimens and 33% of the County list. For the record, my 2019 VC55 Year List is:

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Agelena labyrinthica
Amaurobius fenestralis
Amaurobius similis
Araneus diadematus
Araneus quadratus
Araniella cucurbitina sensu stricto
Bathyphantes gracilis
Cheiracanthium erraticum
Clubiona brevipes
Clubiona comta
Clubiona corticalis
Clubiona lutescens
Clubiona pallidula
Clubiona phragmitis
Clubiona reclusa
Clubiona terrestris
Coelotes atropos
Cyclosa conica
Diaea dorsata
Dictyna uncinata
Dicymbium nigrum/brevisetosum agg.
Diplocephalus latifrons
Diplostyla concolor
Dismodicus bifrons
Drapetisca socialis
Enoplognatha latimana
Enoplognatha ovata
Entelecara acuminata
Episinus angulatus
Erigone atra
Erigone dentipalpis
Ero aphana
Ero furcata
Evarcha falcata
Floronia bucculenta
Gnathonarium dentatum
Gonatium rubellum
Gongylidium rufipes
Harpactea hombergi
Heliophanus flavipes
Hylyphantes graminicola
Kaestneria pullata
Larinioides cornutus
Lathys humilis
Lepthyphantes minutus
Linyphia hortensis
Linyphia triangularis
Maso sundevalli
Metellina mengei
Metellina segmentata
Micrargus herbigradus
Microlinyphia pusilla
Microneta viaria
Monocephalus fuscipes
Neottiura bimaculata
Neriene clathrata
Neriene montana
Neriene peltata
Nigma walckenaeri
Nuctenea umbratica
Oedothorax fuscus
Pachygnatha clercki
Pachygnatha degeeri
Paidiscura pallens
Parasteatoda lunata
Parasteatoda simulans
Pardosa amentata
Pardosa saltans
Philodromus albidus
Philodromus aureolus
Philodromus cespitum
Philodromus dispar
Pholcus phalangioides
Phylloneta impressa
Pisaura mirabilis
Platnickina tincta
Pocadicnemis juncea
Porrhomma microphthalmum
Porrhomma pygmaeum
Pseudeuophrys lanigera
Psilochorus simoni
Savignia frontata
Scotophaeus blackwalli
Selimus vittatus
Steatoda bipunctata
Steatoda grossa
Stemonyphantes lineatus
Tegenaria duellica
Tenuiphantes flavipes
Tenuiphantes tenuis
Tenuiphantes zimmermanni
Tetragnatha extensa
Tetragnatha montana
Tetragnatha obtusa
Theridion mystaceum
Theridion pictum
Theridion varians
Tibellus oblongus
Uloborus plumipes
Walckenaeria acuminata
Walckenaeria unicornis
Xysticus cristatus
Xysticus ulmi
Zygiella x-notata

2019 - My Year in Moths

I think of 2019 as a poor year for moths, but the numbers don't support that conclusion. In 2018 I trapped 1646 moths of 160 species in 53 sessions in my suburban garden. In total I trapped for 469 hours catching 3.51 moths/h. In 2019 I trapped 1725 moths of 145 species in 44 sessions. In total I trapped for 367 hours catching 4.70 moths/h. My trapping effort was reduced, partly by other personal commitments earlier in the year and partly by the weather from mid-September onwards. 2019 in a nutshell - more moths but less diversity.

Top 12:

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The standout on this list for me is Dark Arches. I have never seen so many and they are my abiding memory of 2019.


Previous years:

2018: https://ajcarthropoda.blogspot.com/2019/01/2018-year-in-moths.html
2017: https://ajcarthropoda.blogspot.com/2017/12/2017-year-in-moths.html

"Can you recommend a microscope?"

I am not an expert microscopist and this is only my personal opinion. If your opinion is different to mine, good for you. However I keep getting asked "What microscope do you use? and "What microscope should I buy?", so rather than writing out the answer every time, here are my thoughts.

At some point every aspiring entomologist realises that they need a microscope. Some time later, they realize that they need a better microscope .... and so it goes. The sad fact is with microscopes, you pretty much get what you pay for (unless you are able to find a used microscope in good condition). Buying a cheap microscope may turn out to be a waste of money, buying a very cheap microscope definitely will. However, it would be stupid to spend £10,000 on a microscope if you've never used one before.

"What microscope do you use?"
I currently own three microscopes, one compound and two stereomicroscopes. In essence, a compound microscope will give higher magnifications than a stereomicroscope (there are other important differences but I'm not going into the technicalities here). 95% of my microscopy is done using a stereomicroscope but I find that I also need a compound microscope or a regular basis.



My compound microscope is an Apex Practitioner which I bought from Amazon. In recent years there has been a flood of Chinese optics into both the camera and the microscope markets. The very cheapest stuff is crap, but there are some outstanding examples of good value for money, and the Apex Practitioner is one. You can buy better microscopes if you pay more, but I would highly recommend this microscope if you're starting out.

My first stereomicroscope was an Apex Explorer, also bought from Amazon. It's not bad value for money, but after a few months I realized that I needed a better stereomicroscope. I have kept the Apex Explorer for several reasons - it's light, compact and easy to transport, and if I ever needed to I could run it off a battery pack - it's a microscope you could use in the field or take on holiday with you if you wanted to.



After a while I upgraded to a GXM XTL3T101LED trinocular stereo zoom microscope which I bought from OneStopNature. You may be able to find it cheaper elsewhere but you may end up paying more in the long run. I would strongly recommend phoning Richard Campey for a chat about what you want to do with a microscope before you buy anything.



"So that's all there is to it?"
The advantage of a trinocular microscope is that you can couple a camera directly to it rather than fiddle about trying to take photos through the eyepieces. The type of camera largely doesn't matter (for the record, I use a Sony a7RII), but coupling a camera to microscope is not entirely straightforward as additional optics are involved. The lighting you use is more important than the camera. I mostly use a cheap LED ring light but for some subjects I also use the illumination built into the XTL3T and/or additional Ikea Jansjö LED lamps. Diffusion of the light source is important or the image quality will be poor, and since microscope depth of field is very shallow, image stacking and post-processing is needed to get a clear image of a whole insect.

There is one final and important caveat: a microscope is only as good as the person using it. An expensive microscope won't make you a good microscopist. The key thing is experience. If you are able to spend time learning from an experienced microscopist, this will accelerate your progress.

Good luck!

Imagine My Surprise

I've been chasing Philodromus albidus for the last couple of years, certain that they're out there. VC55 is on the north west edge of their historical range, and there is only one previous record for the vice county. In May 1994 Jon Daws collected a specimen from Essendine Churchyard, just yards inside the county boundary, which John Crocker identified as Philodromus rufus. When Segers redescribed P. albidus in 1989 (Segers, H. (1989) A redescription of Philodromus albidus Kulczynski, 1911 (Araneae, Philodromidae) Bull.Br.arachnol.Soc. 8(2): 38-40), Jon Daws changed this record to P. albidus.

In the last couple of years I've had a number of specimens from my own garden I was pretty sure were P. albidus but they were all immature and so this could not be confirmed. I've been taking advantage of the lousy weather for a mad end of year dash through my backlog of specimens and was delighted to find not one but two adult female Philodromus albidus beaten from Oak at Narborough Bog on 28.06.2019. At last, proof that albidus is out there and that VC55 is a stop over on the north westwards march of this species.

Let There Be Light - Why it's harder to be an entomologist than a botanist!

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I recently treated myself to a new piece of kit. I've lost count of the number of hand lenses I have - unlike some people, I don't lose them on a regular basis (never buy a hand lens without a lanyard) - I just like hand lenses! Being a botantist is so easy, you can just hold a specimen up to the light and look at it with your hand lens (sorry)! Inevitably when I'm looking at tiny bugs I'm always working in my own shadow. My new gizmo is a 10X LED Triplet Loupe Hand Lens. It's a good quality (triplet) hand lens with a tiny LED ring light built in operated by a tiny switch (arrow). It's the same size and weight as a regular hand lens but two or three times the price. I haven't had the opportunity to use it much yet, but I know it's going to be good. I took a quick test shot of a Pine Ladybird (Exochomus quadripustulatus) just by holding my mobile phone to the lens. Inevitably a shiny subject such as this gives a reflection, but this was taken in a dark corner with no additional lighting, so I'm pleased with it. I don't like the fact that it has batteries to replace but given that the lights are LED that shouldn't be too often. Overall, a very good piece of kit which I would recommend.

eDNA is here, it's just not evenly distributed

I worked with DNA for decades (although not strictly speaking eDNA - Environmental DNA), but earlier this year a paper was published that changed my world view (Environmental DNA metabarcoding of wild flowers reveals diverse communities of terrestrial arthropods. (2019) Ecology and Evolution, 9(4), 1665-1679 https://doi.org/10.1002/ece3.4809). I am in no doubt that in a generation's time, eDNA will have changed the way biological recording works, but what I struggle to see is what this will mean for the amateur naturalist.

Consequently, I was interested to come across LifeScanner - "LifeScanner employs DNA barcoding technology to identify insects, food, fur, or any animal tissue. Find out what you're eating, what's in your garden, or in your home." As far as I can make out, the service is only available in the USA & Canada. It's not clear to me exactly what range of taxa the service covers. It doesn't do plants but I'm not clear what arthropods, if any, are covered. Leaving that aside, the price is US$40 for four tests. Would I use a service like this for difficult spiders or springtails which can only identified by their DNA? Possibly, but at 10 quid a pop, not very often.

In the UK the closest thing we currently have is the NatureMetrics Great Crested Newt eDNA service. I'm not aware of any other UK consumer-level eDNA services, but if there are any, please let me know. PCRing your newt soup will cost you £170+VAT, well worth while if you don't want your garden turned into a supermarket car park. NatureMetrics also run a citizen science eDNA project, although I can't figure out exactly what this entails. So it seems that UK arthropod consumer level eDNA on demand is still some way away, but probably not too far. After that, it's all about commoditization and price reduction. And what that will do to field and identification skills is an open question...

Christmas has come early!

On Sunday evening I was going to bed, walked into the kitchen, turned the light on and found ... something crawling across the worktop. At first I thought it was a tiny harvestman, but since that didn't make any sense (it was late!), it had to be a spider. I presumed it must be Pholcus phalangioides but I popped it in a pot and went to bed. Looking at it in the morning it turned out to be a male Psilochorus simoni, although it took me a while to convince myself of that. This species is thought to be of American origin, imported first into France and then into Britain on wine bottles. There are only two previous VC55 records, one from the wine cellar of a very grand country house. I would like to point out that my wine cellar consists of a few bottles from Sainsbury's in the understairs cupboard, and that alone is pretty good evidence that this species is increasing in frequency. The only reason we don't have more records is because it is so easily confused with Pholcus phalangioides (he wrote, without any sense of irony.... ).

Oligolophus hanseni

Oligolophus hanseni is probably not uncommon, but it is difficult to identify, being very similar to Oligolophus tridens. Identification requires either examination with a very good hand lens or a microscope to see the that the trident and associated tubercles are different. Although the trident members of both species are quite small, in Oligolophus hanseni they are normally pale and on close inspection, tipped with black spines. In Oligolophus tridens the trident tubercles can be pale, or the same colour as the body. Oligolophus tridens can have no additional tubercles to the side of the trident and as few as two behind, but in Oligolophus hanseni there are normally more than this to the side and behind. In Oligolophus tridens the two rows of tubercles on the ocularium are often neatly delineated in white, giving the impression of two white eye-rings. In Oligolophus hanseni there are white patches on the ocularium, but these don't usually give the impression of two neat eye-rings. There are only seven previous records for VC55, but in reality I suspect this is quite a common species.

Benchmarking Spider Recording

Recently, I've been thinking a lot about occupancy models for invertebrates (see: Filling the White Holes). Other taxa, notably birds and butterflies (through the BTO Wetland Bird Survey and Butterfly Conservation's UK Butterfly Monitoring Scheme (UKBMS), respectively) have good negative data, i.e. an indication of where species are absent as well as where they are present. For most invertebrate taxa, partly because of lack of resource (recording effort) but mostly because of the inefficiency of recording, all we have are "White Holes" - gaps in the data which are difficult to interpret. The BAS Spider Recording Scheme does not include "negative data" because it's virtually impossible (without DNA approaches) to be certain a spider is truly absent from a particular area. This makes occupancy models difficult if not impossible to derive. The alternative is to fall back to benchmark species as indicators of recording coverage.

Following Filling the White Holes I had an interesting online chat with Geoffrey Hall who introduced me to the idea of axiophytes (it's a botany thing). BSBI gives the following criteria for axiophytes:

• 90% restricted to these conservation habitats
• Recorded in fewer than 25% of tetrads in the county

It seems that Pliny made it up (as so many other things) in his "Natural History". As far as I can tell, neither axioentomos nor axioarachnos exist, so I've just made up two new words (take that Pliny). The point is that axiowhatevers focus attention on presence (even if it is rare presence), rather than absence, so I still tend towards the idea of using a "universal" benchmark species rather than an axiomatic one. Clearly, the choice of a benchmark species is crucial. For springtails, I am happy to use Orchesella cincta, widely acknowledged to be the commonest species of springtail in lowland Britain. I would be amazed if this species was not present in every quadrat in VC55, it's identifiable in the field with no need to put it under a microscope. For spiders it's a little more complex. The obvious choice is Araneus diadematus. While this might seems to be the most obvious choice, it's only the 4th most commonly recorded species in VC55 (n=724) with only 39% of the count of the most frequently recorded species (Tenuiphantes tenuis, n=1,849). I'm pretty sure that this does not reflect the actual situation, but rather recording bias/snobbery (of which I am probably guilty). While the ubiquity of this species is a good reason to think that this is a valid choice, I needed to test the hypothesis. As a starting point I used quadrat mapping - arbitrarily dividing VC55 into a grid and looking at the number of records within each section. A 25x25 grid worked but the the intervals were a bit small and a 10x10 grid is more informative (all VC55 Spider records to end 2018):


The grid for Araneus diadematus looks like this:


To make sense of this, I converted the distributions into histograms:


The distributions look look similar, but to be sure, I ran some further analysis:


While there is a correlation between the Araneus diadematus distribution and the overall VC55 spider records dataset and this is statistically significant (p = 4.49e-11), it's not a great fit (R² = 0.48). In contrast, when I run the same exercise on the benchmark species I use for normalizing springtail recording effort (Orchesella cincta), I get an R² value of 0.87 (p = 2.2e-16), so that's a much better fit from a dataset which is nearly ten times smaller than the VC55 spider data. So I conclude that Araneus diadematus is a valid benchmark for VC55 spider recording - but it's not a great one. If you can think of a better candidate benchmark species, please let me know.


Acknowledgements:
All data Copyright Leicestershire and Rutland Environmental Records Centre.
Data visualization performed using the R platform, v. 3.6.1 (R Core Team (2014) R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. http://www.R-project.org).
J. Cann for assistance with data visualization.

Rainbow Dust [Book Review]

Rainbow Dust: Three Centuries of Delight in British Butterflies
Peter Marren
Vintage, 2016
ISBN: 1784703184

Cover blurb: "Rainbow Dust explores the ways in which butterflies delight and inspire us all, naturalists and non-naturalists alike. Beginning with the author's own experience of hunting and rearing butterflies as a boy, Peter Marren considers the special place of the butterfly in art, literature, advertising and science, and, latterly, our attempts to conserve them. Rainbow Dust takes in the controversy over collecting, the women who studied them and the curious details that lead to butterflies being feared as well as loved. This is a celebration of butterflies; one shot through with a sense of wonder but also of sorrow at what we are losing."

Peter Marren is now a highly respected nature writer best known for the excellent Bugs Britannia and Chasing the Ghost: My Search for all the Wild Flowers of Britain. Although I enjoyed reading Rainbow Dust and learned a lot from it, this is very much in the old school of nature writing, and personally, I enjoyed The Moth Snowstorm more and would recommend that over this book. Nevertheless, Rainbow Dust is still a good read and I recommend it.